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Abstract

Here, we describe a sorbitol-based optical clearing method, called modified Sca/eS that can be used to image all hair cells (HCs) in the mouse cochlea. This modification of Sca/eS is defined by three steps: decalcification, de-lipidation, and refractive index matching, which can all be completed within 72 h. Furthermore, we established automated analysis programs that perform machine learning-based pattern recognition. These programs generate 1) a linearized image of HCs, 2) the coordinates of HCs, 3) a holocochleogram, and 4) clusters of HC loss. In summary, a novel approach that integrates modified Sca/eS and programs based on machine learning facilitates quantitative and comprehensive analysis of the physiological and pathological properties of all HCs.

Sound waves reach the inner ear via conduction mechanisms in the external and middle ear. The cochlea is an organ in the inner ear that transduces mechanical stimulation into electronic signals. As a result, generated auditory information is transferred to the brain. Hair cells (HCs) in the organ of Corti, an auditory sensory epithelium, are protected by the temporal bone and are precisely disposed to detect every frequency band from the base (high pitch) to the apex (low pitch) (von Békésy, 1990). Functional analysis of the cochlea is limited due to its anatomical specificity. Surface preparation and sectioning (paraffin or frozen sections) are the most popular methods in histological analysis; however, these techniques do not preserve the three-dimensional structure (Fujimoto et al., 2017; Mizushima et al., 2017). To better understand the processing of auditory information, the location and diversity of HCs must be accurately determined. Consequently, a novel technique to identify intact HCs has long been sought. Established optical clearing methods such as CLARITY, 3DISCO, Sca/eS, iDISCO, and CUBIC (Dodt et al., 2007; Chung et al., 2013; Renier et al., 2014; Susaki et al., 2014; Hama et al., 2015) enable imaging of intact brains and other organs, but are not optimized for imaging of all HCs (Urata et al., 2019). Based on recently developed optical clearing methods for hard tissues such as vDISCO, Bone CLARITY, PEGASOS, CUBIC-X, PACT, and PARS (Calve et al., 2015; Treweek et al., 2015; Berke et al., 2016; Greenbaum et al., 2017; Cai et al., 2018; Jing et al., 2018; Tainaka et al., 2018), we present a sorbitol-based optical clearing method (modified Sca/eS) that is optimized for intact imaging of all mouse HCs in the cochlea (Urata et al., 2019).

By acquiring successive images of HCs using modified Sca/eS, we established a novel algorithm for comprehensive analysis of HCs. These programs are available on GitHub (https://github.com/okabe-lab/cochlea-analyzer.git). Use of modified Sca/eS in combination with these programs facilitates quantitative and comprehensive analysis of the physiological and pathological properties of all HCs.

Figure 1. Time course of modified Sca/eS. The inner ear embedded in the temporal bone was removed (panel A) and dissected along the fistula (B) (dotted lines in the left of panel A). The cochlea was separated (dotted line in panel C) from the semicircular canals along the fissure (red arrow in panel C). The cochlea was removed [panel D and (i)] and decalcified (ii). The sample was cleared immediately after being submerged in solution 1 (iii). The sample was then submerged in solution 2 and its transparency gradually increased (iv-viii). A: anterior, P: posterior, M: medial, L: lateral, C: cochlea, SC: semicircular canal, RI: refractive index. Scale bar: 1 mm.

Extraction of the cochlea from the temporal bone

Fix the cochlea in 4% paraformaldehyde prepared in PBS overnight at 4 °C with gentle shaking.

Wash the cochlea three times (15 min each) with PBS.

Incubate the samples in 500 mM EDTA prepared in PBS for 48-120 h at 37 °C.Note: The incubation duration depends on the sample size. For example, 48 h for pups, 72 h for young (2-week-old) mice, and 120 h for middle-age (6-month-old) mice.

Incubate the samples in a solution containing the appropriate dilutions of primary antibodies for 2-48 h at 37 °C.

Wash the samples three times (30 min each) with PBS containing 0.1% Triton X-100 with continuous rocking at 40 rpm.

Detect primary antibodies by incubating the samples with a solution containing secondary antibodies for 12-48 h at 37 °C followed by three washes with PBS containing 0.1% Triton X-100.

Sample preparation for imaging (see Figures 2 and 3)

Roll up a piece of Blu Tack into a cylindrical shape that is slightly thicker than the cochlea [Figure 2 (i)].

Align Basukoku in a horseshoe shape on the glass slide [Figure 2 (ii)].

Place the Blu Tack on the Basukoku [Figure 2 (iii, iv)].

Place a drop of Basukoku inside the Blu Tack horseshoe to make a pedestal for the cochlea [Figure 2 (v)].

Carefully place the cochlea on the pedestal [Figure 2 (vi)].Note: Position the cochlea so that the axis of the center of the cochlea is aligned perpendicular to the glass slide (Figure 2).

Place the cover glass on the surface of the Blu Tack and gently push it toward the surface of the cochlea [Figure 2 (vii)].Note: Keep pushing the cover glass until it reaches the surface of the cochlea. This process is critical to image the whole cochlea [Figure 2 (vii-a)].

With the horseshoe opening facing upward, add 100 μl of solution 2 (see Recipes) to the gap within the horseshoe until the imaging chamber is filled [Figure 2 (viii)].Note: Images iv-viii in Figure 1 show how transparent the cochlea becomes after 20 min (images were acquired at increments of 5 min).

Fill the gap between the Blu Tack with Basukoku to seal the horseshoe opening [Figure 2 (ix)].Note: Air bubbles should not remain in the chamber; this would indicate potential leaking of solution 2 and cause the sample to dry out.

Figure 2. Sample preparation for the imaging step

Figure 3. Cochlea position. The center of the first (green) and second (red) turns of the cochlea. Images on the left are the ideal position for imaging of HCs in the whole cochlea, where d represents the distance between the centers of the green and red areas. Upper panels (A) show all HCs imaged using a two-photon microscope in a successful case. Lower panels (B) are representative cases where the approach failed. Images on the left are from the top view, while images on the right are from the side view. The modiolus of the cochlea should be perpendicular to the slide glass [i.e., centers should overlap (purple)].

Imaging of the sample

Place one drop of distilled water on the cover glass and then place the objective lens against the cover glass.Note: Use an appropriate immersion media for high-resolution imaging. Water-immersion lenses perform much better than air-immersion objective lenses.

To image the whole cochlea, images should be successively acquired with 10-40% overlap [Figure 2 (i)].

See example images of sample processed with previous well known clearing methods (Figure 4).

Figure 4. Comparison with other clearing methods. MYO7A-immunopositive HCs in samples were not detected by 3DISCO, iDISCO, CLARITY, or CUBIC (A). Microdissection of the membrane labyrinth of the iDISCO-processed samples in panel A confirmed the presence of MYO7A-immunopositive HCs, suggesting that the surrounding bone tissue prevented the detection of fluorescence (B). Scale bars: 500 µm in panel A and upper image of panel B, 100 µm in lower image of panel B.

Automated HC analysis protocol based on machine learning
HCs were automatically detected and analyzed using custom MATLAB scripts (R2017b, MathWorks), the details of which were provided in a previous study (Urata et al., 2019). The MATLAB source code is available on GitHub (https://github.com/okabe-lab/cochlea-analyzer.git). Data analysis procedures including statistical analysis were described in the original publication of the protocol (Urata et al., 2019). The actual running of the program was shown in this protocol paper.

The main0.m program (1 min) checks the integrity of the software so that automated analysis can be flawlessly performed (see Video 1).Note: If an error occurs during the process, double check to see if any required files are absent under “Function Files” on GitHub (https://github.com/okabe-lab/cochlea-analyzer.git).

Video 1. Checking the integrity of the software before performing automated analysis using main0.m

The main1.m program merges all the files together to create a linearized image of HCs, which takes about 15 min in total (Step B2 in Figure 5 and see Video 2).Note: If the main0.m program completes without an error, it is guaranteed to function properly. A TIFF image is produced in this step.

Video 2. Merging of all the acquired files together to create a linearized image of HCs using main1.m

Figure 5. Image generation by the programs. Raw images are stitched (Step 1), and the lines between inner and outer HCs are illustrated (orange line in Step 2). Finally, HCs are linearized and representative images of the apical, middle, and basal portions are enlarged. Colored dots in Step 3 and Step 4 indicate the presence of HCs, and asterisks represent loss of HCs.

Solution 2 (20 ml)
5.7 g of guanidinium chloride (or 4.8 g of urea)
12 g of D-sorbitol
20 µl of Triton X-100 in PBS (pH 7.1)Note: The solution can be kept at room temperature for up to 2 months for good clearing.

We thank Dr. Shigeo Okabe for helpful suggestions and supervision. This method was adapted from our original publication of the protocol (Urata et al., 2019). This work was supported by grants from the Japan Society for the Promotion of Science KAKENHI (15K10743, 2653081, and 16K15717), the Japan Science and Technology Agency (JPMJCR14W2), the Japan Agency for Medical Research and Development (17gm5010003), the Japan Society for the Promotion of Science (17H01387), the UTokyo Center for Integrative Science of Human Behavior, and the Ministry of Education, Culture, Sports, Science, and Technology (18H04727 and 26111506).